Metabolism of the insecticide metofluthrin in cabbage (Brassica oleracea)

Article abstract of DOI:10.1021/jf203903r

The metabolic fate of metofluthrin [2,3,5,6-tetrafluoro-4-(methoxymethyl) benzyl (E,Z)-(1R,3R)-2,2-dimethyl-3-(prop-1-enyl)cyclopropanecarboxylate] separately labeled with ¹⁴C at the carbonyl carbon and the α-position of the 4-methoxymethylbenz

Full text of DOI:10.1021/jf203903r

Article
pubs.acs.org/JAFC
Metabolism of the Insecticide Metofluthrin in Cabbage (Brassica
oleracea)
Daisuke Ando,* Masao Fukushima, Takuo Fujisawa, and Toshiyuki Katagi
Environmental Health Science Laboratory, Sumitomo Chemical Company, Ltd., 4-2-1 Takatsukasa, Takarazuka, Hyogo 665-8555,
Japan
ABSTRACT: The metabolic fate of metofluthrin [2,3,5,6-tetrafluoro-4-(methoxymethyl)benzyl (E,Z)-(1R,3R)-2,2-dimethyl-3-
(prop-1-enyl)cyclopropanecarboxylate] separately labeled with 14C at the carbonyl carbon and the α-position of the 4-
methoxymethylbenzyl ring was studied in cabbage (Brassica oleracea). An acetonitrile solution of 14C-metofluthrin at 431 g ai
ha−1 was once applied topically to cabbage leaves at head-forming stage, and the plants were grown for up to 14 days. Each
isomer of metofluthrin applied onto the leaf surface rapidly volatilized into the air and was scarcely translocated to the untreated
portion. On the leaf surface, metofluthrin was primarily degraded through ozonolysis of the propenyl side chain to produce the
secondary ozonide, which further decomposed to the corresponding aldehyde and carboxylic acid derivatives. In the leaf tissues,
the 1R-trans-Z isomer was mainly metabolized to its dihydrodiol derivative probably via an epoxy intermediate followed by
saccharide conjugation in parallel with the ester cleavage, whereas no specific metabolite was dominant for the 1R-trans-E isomer.
Isomerization of metofluthrin at the cyclopropyl ring was negligible for both isomers. In this study, the chemical structure of each
secondary ozonide derivative was fully elucidated by the various modes of liquid chromatography−mass spectrometry (LC-MS)
and nuclear magnetic resonance (NMR) spectroscopy together with cochromatography with the synthetic standard, and their
cis/trans configuration was examined by the nuclear Overhauser effect (NOE) difference NMR spectrum.
KEYWORDS: metofluthrin, plant metabolism, ozonide, diastereomer
the prop-1-enyl group and the benzyl carbon to form
dicarboxylic acid (11) and terephthalic acid (9) derivatives,
respectively.2 Further degradation of the metabolites resulted in
production of carbon dioxide, but little enantiomerization and
geometrical isomerization proceeded. Radioactivity was also
detected as the soil bound residues. The soil adsorption
coefficient (Koc) of 1-RTZ in three German soils was
determined to be 3553−6142 mL g−1 oc by the batch
equilibrium method.2 Under illumination on the moisture-
controlled soil, 1-RTZ degraded with a half-life of 8.1−12.0
days.3,4 The photodegradation pathway was oxidation of the
double bond at the prop-1-enyl moiety and cleavage of the ester
linkage, followed by further decomposition to polar compounds
and mineralization to carbon dioxide. The major degradates
detected were carbonaldehyde (4) and carboxylic acid (5)
derivatives, which were decomposed from ozonide (2) and diol
(6) produced by the activated oxygen species, that is, ozone
and hydrogen peroxide, respectively. The photoisomerization
hardly proceeded throughout the test duration.3 In the
hydrolysis study, 1-RTZ was moderately degraded by ester
cleavage with a half-life of 26.8 days at pH 9 and 25 °C, whereas
it was stable at pH 5 and 7. For aqueous photolysis using a
xenon arc lamp, 1-RTZ and 1-RTE rapidly dissipated with half-
lives of 1.1−3.4 days at pH 7 mainly due to ester cleavage to
produce 7 and 10 and oxidation of the olefinic double bond to
form 3 and 4, whereas isomerization was a minor route.4
INTRODUCTION
■
Metofluthrin (1) [SumiOne, Eminence; 2,3,5,6-tetrafluoro-4-
(methoxymethyl)benzyl (E,Z)-(1R,3R)-2,2-dimethyl-3-(prop-
1-enyl)cyclopropanecarboxylate] is a pyrethroid insecticide
for household and public hygiene usages developed by
Sumitomo Chemical Co., Ltd., and has a strong knockdown
activity, especially against mosquitoes. Because of its high
volatility (1.96 × 10−3 Pa) with a low mammalian toxicity, 1 can
be used in nonheated formulations such as fan-type, paper, and
resin emanators.1 Incidentally, the presence of the two optical
centers at the cyclopropenyl ring and one geometrical
isomerism at the propenyl side chain result in eight isomers,
and 1 consists of the biologically active 1R-trans isomers having
an E/Z geometrical ratio of 1/8, abbreviated as 1-RTE and 1-
RTZ.1
The extremely limited emission of 1 to the environment is
most likely because it is mainly used indoors. However, as there
is no geographic restriction of its use, that is, personal outside
insect repellent, 1 may directly or indirectly reach a nontarget
environment such as a water body, soil, and plants. In addition,
with regard to biocide, submission of relevant data to the
European Union and the U.S. Environmental Protection
Agency regarding environmental fate has recently become
inevitable, irrespective of the use pattern especially due to the
possible contamination by sewage treatment plant effluent.
From these aspects, it is important to obtain experimental data
on 1 to show that it is benign to the environment. The aerobic
soil metabolism in two U.S. soils has shown that 1-RTZ and 1-
RTE rapidly degrade at similar half-lives of 2.3−3.5 days via
cleavage of the ester linkage to produce the corresponding
alcohol (10) and acid (7), followed by successive oxidation at
Received: September 26, 2011
Revised: December 21, 2011
Accepted: December 26, 2011
Published: December 26, 2011
© 2011 American Chemical Society
2607
dx.doi.org/10.1021/jf203903r | J. Agric. Food Chem. 2012, 60, 2607−2616

Journal of Agricultural and Food Chemistry
Article
Figure 1. Proposed metabolic pathway of metofluthrin in/on cabbage plant.
thalic acid (9); (1R,3R)-2,2-dimethyl-3-(E,Z)-propenylcyclopropane-
carboxylic acid (10); (1R,3R)-2,2-dimethyl-3-carboxycyclopropanecar-
boxylic acid (11). The chemical purity of each standard was
determined to be >95% by high-performance liquid chromatography
(HPLC). Degradate 2 was prepared according to the following
method. Three milligrams of either 1-RTZ or 1-RTE was dissolved in 1
mL of n-hexane solution and cooled to < −40 °C using dry ice/
acetone prior to the ozone oxidation. Ozone gas produced at an
approximate concentration of 20 g N−1m−3 using an ozone generator
(type 0N-1-2, Nippon Ozone Co., Ltd., Japan) was gently bubbled
into the reaction mixture for 3 min at < −40 °C, which resulted in the
formation of a white precipitate of crude 2. After the solution returned
to room temperature, 2 was successively purified by HPLC, and the
solvent was removed using an evaporator and dried in vacuo to obtain
a colorless liquid, the chemical purity of which was determined to be
>95%. The chemical structure of 2 was confirmed by various modes of
NMR and LC-ESI-MS spectroscopies. 2 was stable in acetonitrile, n-
hexane, and chloroform for a few months in a freezer below 0 °C, but
rapidly decomposed to 4 and 5 in aqueous organic media. The
following 14C-labeled isomers of 1 were synthesized in our laboratory
with the reported methods;2−4 1-RTZ separately labeled at the α-
position of 2,3,5,6-tetrafluoro-4-methoxymethylbenzyl ring (ben-
zyl-14C) or the carbonyl carbon (carbonyl-14C) and 1-RTE labled at
the carbonyl carbon (carbonyl-14C) with specific activities of 0.1654,
0.1651, and 0.1651 mCi mg−1, respectively. β-Glucosidase (almond)
and cellulase (Aspergillus niger) were purchased from Wako Pure
Chemical Industries, Ltd. All of the reagents and solvents used were of
the analytical grade.
Up to this date, the plant metabolism study of 1, for which
information is necessary for the ecotoxicological assessments of
nontarget species, is not available. From this standpoint, we
have studied the metabolism of 1 in cabbage grown in a
greenhouse, which was selected as a model plant because its
broad and waxy leaves5 are considered to be suitable to trap and
prevent the loss of 1 by rapid volatilization.6 The metabolites
formed in/on the plant were clarified by extensive spectro-
metric analysis using LC-MS and NMR spectroscopy in
conjunction with direct chromatographic comparison with the
synthetic standards.
MATERIALS AND METHODS
■
Chemicals. The nonradiolabeled metofluthrin (1) and its potential
degradates as follows were synthesized in our laboratory according to
the reported methods.2−4 The structures of the compounds are
referred to Figure 1; 2,3,5,6-tetrafluoro-4-(methoxymethyl)benzyl
(1R,3R)-2,2-dimethyl-3-(3-methyl-1,2,4-trioxolane-5-cyclopropanecar-
boxylate (2); 2,3,5,6-tetrafluoro-4-(methoxymethyl)benzyl (1R,3R)-
2,2-dimethyl-3-(3-methyl-1,2-epoxy)cyclopropanecarboxylate (3);
2,3,5,6-tetrafluoro-4-(methoxymethyl)benzyl (1R,3R)-2,2-dimethyl-3-
formylcyclopropanecarboxylate (4); 2,3,5,6-tetrafluoro-4-
(methoxymethyl)benzyl (1R,3R)-2,2-dimethyl-3-carboxycyclopropa-
necarboxylate (5); 2,3,5,6-tetrafluoro-4-(methoxymethyl)benzyl
(1R,3R)-2,2-dimethyl-3-(1,2-propanediol)cyclopropanecarboxylate
(6); 2,3,5,6-tetrafluoro-4-(methoxymethyl)benzyl alcohol (7); 2,3,5,6-
tetrafluoro-4-(methoxymethyl)benzoic acid (8); tetrafluorotereph-
2608
dx.doi.org/10.1021/jf203903r | J. Agric. Food Chem. 2012, 60, 2607−2616

Journal of Agricultural and Food Chemistry
Article
Chromatography. The reversed-phase (RP) HPLC analysis of
metabolites within the leaf surface rinse and extract was conducted
using a Hitachi LC module (model L-7000) equipped with a
SUMIPAX ODS A-212 column (5 μm, 6 mm i.d. × 15 cm, Sumika
Chemical Analysis Service (SCAS), Ltd.) at a flow rate of 1 mL min−1.
The following gradient system was operated as the typical analysis with
acetonitrile containing 0.05% formic acid (solvent A) and distilled
water with 0.05% formic acid (solvent B): 0 min, % A/% B, 5/95; 0−3
min, 5/95, isocratic; 3−10 min, 45/55 at 10 min, linear; 10−70 min,
75/25 at 70 min, linear; 70−71 min, 5/95 at 71 min, linear; 71−80
min, 5/95, isocratic (HPLC method 1). For the separation of four
isomers of 2, the gradient system as follows was applied: 0 min, % A
(acetonitrile)/% B (water), 53/47; 0−3 min, 53/47, isocratic; 3−43.7
min, 58/42 at 43.7 min, linear; 43.7−44 min, 75/25 at 44 min, linear;
44−51 min, 53/47 at 51 min, linear; 51−60 min, 53/47, isocratic
(HPLC method 2). The chiral analysis was conducted with a
Shimadzu LC-10AT HPLC system connecting two SUMIPAX DI-
NO2 columns (5 μm, 4 mm i.d. × 25 cm, SCAS) and one
CHIRALCEL OD-H column (5 μm, 4.6 mm i.d. × 25 cm, Daicel
Chemical Industries, Ltd.) in series using an isocratic eluent of n-
hexane/ethanol, 1000/0.5 (v/v), at a flow rate of 0.9 mL min−1. The
radioactivity eluted was monitored with a Flow Scintillation Analyzer
Radiomatic 500TR (Perkin-Elmer Co., Ltd.) or Ramona (Raytest,
Germany) radiodetector equipped with a 500 μL liquid cell using
Ultima-Flo AP (Perkin-Elmer, Co., Ltd.) as the scintillator. The
detection limit of the HPLC analyses was 30 dpm. The typical
retention times of 1-RTZ and 1-RTE and their related reference
standards have been reported previously.2−4
One- or two-dimensional thin-layer chromatography (1D- or 2D-
TLC) was carried out for an analytical purpose using precoated silica
gel 60F254 thin-layer chromatoplates (20 × 20 cm, 0.25 mm thickness;
E. Merck). The nonradiolabeled reference standards were detected by
exposing the chromatoplates to ultraviolet light or spraying
bromocresol green reagent for direct visualization. Autoradiograms
were prepared by transcribing the TLC plates to BAS-IIIs Fuji imaging
plates (Fuji Photo Film Co., Ltd.) for several hours. The radioactivity
in each spot exposed onto the imaging plate was detected by a Bio-
Imaging Analyzer Typhoon (GE Healthcare). The solvent systems for
2D-TLC were chloroform/methanol, 9/1 (v/v), and toluene/ethyl
acetate/acetic acid, 5/7/1 (v/v/v). For the analysis of 2, n-hexane/
toluene/acetic acid, 3/15/2 (v/v/v) was applied for 1D-TLC
development. The typical Rf values of 1-RTZ and 1-RTE and their
related reference standards have been reported previously.2−4
Spectroscopy. For NMR spectrometric analyses, one-dimensional
(1H, 13C, distorsionless enhancement by polarization transfer (DEPT),
nuclear Overhauser effect (NOE) difference) and two-dimensional
experiments (1H−1H correlation spectroscopy (COSY), heteronuclear
single quantum coherence (HSQC), heteronuclear multiple-bond
connectivity (HMBC), NOE correlated spectroscopy (NOESY)) were
employed in d-chloroform including tetramethylsilane (TMS) using a
Varian Mercury 400 (Varian Technologies Ltd.) spectrometer (400
MHz).
Liquid chromatography−electrospray ionization−mass spectrome-
try (LC-ESI-MS) analysis was conducted using a Waters Micromass
ZQ spectrometer equipped with a Waters separation module 2695 and
photo array detector 2996 as a liquid chromatograph. For the
conventional analysis of metabolites, HPLC method 1 was applied
with the analytical parameters controlled by MassLynx software
(version 4.00) as shown: source temperature, 100 °C; desolvation
temperature, 350 °C; capillary voltage, 3.2 kV; cone voltage, 10−40 V.
For the analysis of 2, conditions of source temperature, 70 °C, and
desolvation temperature, 300 °C, were selected to mitigate its thermal
degradation and the gradient system as follows was applied: 0 min, %
A (acetonitrile)/% B (methanol/20 mM ammonium acetate (20/10,
v/v))/% C (water), 5/30/65; 0−50 min, 20/30/50, linear.
subtracted from the dpm value of a measured sample. The 14C in the
extracted residues and untreated plant portions was converged and
measured as 14CO2 using a Packard model 307 sample oxidizer. 14CO2
produced was absorbed into 9 mL of Packard Carb-CO2 absorber and
mixed with 15 mL of Packard Permafluor scintillator, and the
radioactivity therein was quantified by LSC. The efficiency of
combustion was determined to be >95.8%.
Plant Material and Treatment. Cabbage (Brassica oleracea var.
capitata, cv. Green Ball) was grown in a 1/5000-are Wagner pot filled
with Kasai soil (Hyogo, Japan) in a greenhouse at 25 °C during the
day and at 20 °C during the night. The application of each [14C] -1 to
the cabbage plants was conducted at the plant growth stage of BBCH
41.7 The average surface area and weight of a cabbage leaf at the
application stage were 176.6 cm2 and 10.31 g, respectively. The
characterization of Kasai soil is as described as follows: soil texture
(%), sand 82.9, silt 8.9, clay 8.2; soil classification, sandy loam; organic
carbon content (w/w), 1.7; pH (H2O), 6.6; maximum water-holding
capacity (g per 100 g of dry soil), 28.19.
The application dose of 1 to cabbage plants was 431 g ai ha−1, which
was determined by assuming that a typical commercial can was directly
sprayed onto cabbage leaves as the worst-case scenario: 4 g of aerosol
of the water-based formulation containing 0.1% (w/w) of 1 was
sprayed once onto a square-foot field for the purpose of general insect
repellent. Although the isomeric ratio of 1-RTZ and 1-RTE in the
active ingredient is 8/1, the same dosing rate was used for each isomer
in this study. For the leaf treatment, the dosing solution per leaf was
prepared by mixing 0.167 MBq of each [14C]-1 isomer with 0.761 mg
of the corresponding unlabeled material in 100 μL of acetonitrile
(0.212 MBq mg−1). The prepared dosing solution was topically
applied onto nine leaves (three leaves × three pots) using a
microsyringe for each label. With respect to the soil treatment, the
dosing solution per pot was prepared by combining 1.667 MBq of
each [14C]-1 with 0.862 mg of the unlabeled material in 1 mL of
acetonitrile and was applied onto 120 g of Kasai soil in a plastic bag
using a pipet and thoroughly mixed for 30 min. After evaporation of
acetonitrile, it was gently put onto the soil surface of the Wagner pot
in which cabbage plants were grown.
Sampling, Extraction, and Analysis. For leaf treatment, three
cabbage leaves per pot were harvested at 2, 7, and 14 days after
treatment. The leaves were individually cut from the stem using
scissors, and untreated leaves and stems were similarly sampled with
another uncontaminated one. For soil treatment, both cabbage plant
and soil were separately sampled at 14 days after treatment. The whole
cabbage plant was obtained by cutting the stem just above the ground.
The dried soil was vertically divided into three layers according to its
depth (top, 0−2 cm; middle, 2−10 cm; bottom, 10−18 cm), and the
root was removed from the soil. All samples were immediately
weighed and stored in a freezer (below −20 °C) until analysis.
In the case of the leaf treatment, the surface of treated leaves was
rinsed with 100 mL of methanol per leaf. The rinsed leaf was cut into
small pieces and homogenized with 20 mL of methanol at 10000 rpm
and 0 °C for 10 min using a homogenizer AM-8 (Nissei Ltd., Japan).
The homogenate was vacuum filtered to separate the extract and the
residue. The residue remaining after filtering was extracted again in the
same manner, and the filtrate was combined. The process was repeated
using methanol/water (4/1, v/v). Each aliquot of the surface rinse,
methanol, and methanol/water extracts was analyzed with LSC,
HPLC, and 2D-TLC. The extracted residues were air-dried and
individually combusted for LSC analysis. For the soil treatment, a
portion of each soil layer was subjected to a combustion analysis to
determine the residual amount of 14C. Approximately 50 g of the
evenly mixed top layer soil for each label was transferred into 200 mL
plastic centrifuge bottles, and 100 mL of methanol was added. The
bottle was mechanically shaken for 10 min with a Taiyo SR-IIw
recipro-shaker and then centrifuged at 5000 rpm at 4 °C for 10 min
using a himac CR20G high-speed refrigerated centrifuge (Hitachi Ltd.,
Japan). The extract was recovered from the bottle by decantation, the
residues were repeatedly extracted twice in the same manner, and then
the extracts were combined. The procedure was repeated using
methanol/concentrated HCl (100/1, v/v). After radioassay by LSC,
Radioanalysis. Radioactivity in the liquid surface rinse and extract
from plant was determined by mixing each aliquot with 10 mL of
Packard Emulsifier Scintillator Plus and analyzed by liquid scintillation
counting (LSC) with a Packard model 2900TR spectrometer. The
background level of radioactivity in LSC was 30 dpm, which was
2609
dx.doi.org/10.1021/jf203903r | J. Agric. Food Chem. 2012, 60, 2607−2616

Journal of Agricultural and Food Chemistry
Article
Table 1. Distribution of Radioactivity after Foliar Applications
a
nd, not detected.
Table 2. Distribution of 1 and Its Metabolites in Treated Cabbage Leaves
a
b
c
d
e
nd, not detected; Consisted of multiple components, each of which amounted to <2.0%. Sugar conjugate of 6. −, not available. Consisted of
multiple components, each of which amounted to <7.0%.
the extracts were concentrated using an evaporator, and each 10000
dpm was subjected to HPLC and 2D-TLC analyses. The soil residues
after extraction were air-dried in open vessels at room temperature for
several days and subjected to combustion analysis to determine the
remaining radioactivity.
The total radioactive residue (TRR) in the test system of the foliar
application was determined as the sum of 14C in the surface rinse,
extract and unextractable. The TRR recovered from the test system of
the soil treatment was determined as the sum of 14C in extractable and
unextractable fractions of topsoil and that in the middle and bottom
soil layers and plant.
Identification of Metabolites. In general, the identity of the
metabolites was confirmed by HPLC and 2D-TLC cochromatog-
raphies with the nonradiolabeled reference standards. In addition, LC-
ESI-MS and NMR analyses were carried out for identification of 2. To
identify conjugated metabolites, approximately 200 000 dpm of the
extract was dissolved in 1 mL of 10 mM sodium acetate buffer at pH
5.0, and 10 mg of cellulase or β-glucosidase was added to the solution.
The mixture was incubated in a BR-180LF BioShaker (TAITEC Co.
Ltd., Japan) at 37 °C and 100 rpm. The aliquot of the reaction mixture
was sampled sequentially until 7 days after incubation and directly
analyzed with HPLC and 2D-TLC. With regard to the characterization
of metabolites in the extract, chemical derivatizations (acetylation and
methylation) were performed. The leaf extracts including 100 000 dpm
were taken and the solvent was dried up using an evaporator. The
dried extract was redissolved in 500 μL of pyridine/acetic anhydride
(50/50, v/v) and allowed to stand overnight for acetylation and then
dissolved again in 500 μL of acetonitrile with dropwise addition of 20
μL of 0.1% trimethylsilyldiazomethane in n-hexane and mixing for 4
min for methylation. The derivatized samples were directly analyzed
by HPLC.
For the purpose of collecting a sufficient amount of 2 for LC-MS
and NMR analyses, 0.167 MBq of [benzyl-14C]-1-RTZ mixed with 10
mg of the nonradiolabeled material in acetonitrile was applied onto the
surface of three cabbage leaves. The plants were grown in the
greenhouse for 7 days. After the sampling, the treated leaves were
rinsed with acetonitrile and the rinsate was subjected to isolation of 2
using HPLC method 2.
RESULTS
■
Distribution of 14C in Plant and Soil. The distribution of
radioactivity after the foliar application is summarized in Table
1. For the leaf treatment, the radioactivity recovered from the
treated leaves was 31.4−71.1% of the applied radioactivity (%
AR) after 14 days. The unrecovered 14C was most likely lost by
vaporization, considering the vapor pressure (1.96 × 10−3 Pa)
and Henry’s law constant (2.1 × 10−4 m3 mol−1) of 1. The 14
C
recovered by the surface rinse gradually decreased to 11.8−
26.3%AR at the end of the study, whereas that in the leaf
extract concomitantly increased to 18.0−42.2%AR. The total
2610
dx.doi.org/10.1021/jf203903r | J. Agric. Food Chem. 2012, 60, 2607−2616

Journal of Agricultural and Food Chemistry
Article
Figure 2. RP-HPLC chromatogram of the surface rinse of [carbonyl-14C]-1-RTZ treated leaf after 7 days. Numbers above each peak indicate the
corresponding metabolite.
amount of unextractable 14C was <3.1%AR during the study in
any case. Little radioactivity was translocated from the treated
leaf to the untreated portions of plant. With respect to the soil
treatment, 14C recovered from the test system was 39.6−60.4%
AR after 14 days. The majority of the radioactivity remained in
the top layer of soil (0−2 cm), which amounted to 12.1−30.9%
AR, whereas levels in the middle (2−10 cm) and bottom (10−
18 cm) layers of soil were 13.1−16.1 and 4.8−9.9%AR,
respectively. The uptake of 14C from the soil to the plant was
negligible at 1.5−3.6%AR.
these metabolites are listed in Table 3. Each proton signal
pattern was very similar between the two isomers, but was
clearly different from that of 1-RTZ as its two methine protons
at the propenyl moiety shifted from 4.92 and 5.35 ppm to a
lower magnetic field of 5.11 and 5.62 ppm, respectively. With
1
regard to the H−1H COSY spectrum, no cross signal was
observed between the above two adjacent methine protons for
2a and 2b, which suggested the cleavage of the C−C double
bond (Figure 3). In the 1D-TLC analysis using an acidic
solvent C, both 2a and 2b decomposed to carbonaldehyde (4)
and carboxylic acid (5) derivatives during the development.
Therefore, the chemical structures of 2a and 2b were assumed
to be ozonide derivatives possessing a 1,2,4-trioxolane ring. To
definitively identify the chemical structure of ozonide, 2 was
synthesized from 1-RTZ and 1-RTE. The modification of
HPLC gradient system (method 2) enabled the synthetic 2 to
be separated into four isomers, namely, 2a−2d. The observed
ratio of the four isomers was 2a/2b/2c/2d = 33/16/37/14 for
The metabolite distribution for the leaf treatments is
summarized in Table 2. The representative HPLC chromato-
gram of the surface rinse fraction is shown in Figure 2. On the
leaf surface, 1-RTZ and 1-RTE were rapidly dissipated to <0.1%
of the total radioactive residue (%TRR) at 14 days after
treatment. 2, 4, and 5 were detected as major metabolites, with
maximum amounts reaching 25.4, 28.5, and 26.0%TRR,
respectively, at 7 days after treatment. In addition, 2 was
clearly separated into four peaks by using HPLC method 2; that
is, the retention times of 2a, 2b, 2c, and 2d were 33.7, 31.7,
30.0, and 29.3 min, respectively, which indicated the presence
of four isomers. Other minor metabolites were <2.0%TRR. In
the leaf extract, 1-RTZ and 1-RTE were detected at 0.4−18.6
and 9.4−18.5%TRR, respectively, during the test duration.
Comparison of the metabolic profile has shown that it is
relatively different between 1-RTZ and 1-RTE. In the case of 1-
RTZ, the major metabolite having an intact ester linkage was
the glucose conjugate of 6, amounting to its maximum 12.3%
TRR. 7 was detected at 12.2%TRR for the benzyl label, but no
corresponding degradates unique to the cyclopropane label
exceeding 10%TRR were observed, which suggested faster
degradation for the acid moiety in the cabbage plant. For 1-
RTE, 4 and 5 were formed at 4.0 and 4.6%TRR, respectively,
and none of the metabolites exceeded 7%TRR. To evaluate the
isomerization ratio, the chiral HPLC analysis of 1-RTZ and 1-
RTE remaining on/in the leaf for the benzyl label was
conducted, which demonstrated negligible isomerization to
the other stereoisomers (<0.7%TRR). For the soil treatment,
most of the radioactivity remained as the intact parent
compound except for [benzyl-14C] 1-RTZ, which produced 8
in 41.9%TRR at 14 days after treatment.
1
1-RTZ and 2a/2b/2c/2d = 40/21/28/11 for 1-RTE. The H
and 13C NMR spectra of each isomer are listed in Table 3. The
overall trends of the spectra were similar within four isomers,
but the chemical shifts related to protons H5 and H6 on the
cyclopropane ring were clearly different. The four isomers all
exhibited identical cross peaks in various modes of 2D-NMR
analyses, which clearly demonstrated the preservation of the
chemical structures of the parent compound except for the
cleavage of the C−C double bond at the prop-1-enyl moiety. In
the LC-ESI-MS analysis, the pseudo ions of 2a−2d were
observed at m/z 409 [M + H]+, 426 [M + NH4]+, and 431 [M
+ Na]+ in the positive ion mode, which indicated the addition
of three oxygen atoms to 1. Taking all of the above into
account, these four products were determined to be the
diastereomers of the ozonide derivative, for which the two
asymmetric carbons exist in the trioxolane ring. Incidentally, by
1
1
comparison of the H and H−1H NMR spectra and TLC, the
HPLC cochromatogram has clarified that 2a−2d produced on
the leaf surface were identical with those of the synthesized
standards. In addition, the HPLC-purified synthetic isomers 2a,
2b, and the mixture of 2c and 2d were subjected to NOE
difference analysis to obtain steric information. The NOE
resonance was clearly observed between the protons H10
(5.34−5.36 ppm) and H11 (4.92−4.96 ppm) in the trioxolane
ring for the mixture (2c + 2d), whereas scarce or no resonance
was observed for 2a and 2b, which indicated the structural
distance between these protons is closer for the former isomers
compared to the latter (Figure 4), thus characterizing their
configurations as trans (threo) and cis (erythro), respectively.
Identification of Metabolite 2. Approximately 180 and
250 ng of isomeric metabolites 2a and 2b were obtained,
respectively, from the experiment on collecting sufficient
amounts for their identification. The purified metabolites 2a
1
1
and 2b were individually subjected to H NMR and H−1H
COSY and 1D- and 2D-TLC analyses. The chemical shifts of
2611
dx.doi.org/10.1021/jf203903r | J. Agric. Food Chem. 2012, 60, 2607−2616

Journal of Agricultural and Food Chemistry
Article
1
Table 3. H and 13C NMR Chemical Shifts of 2 Isomers (2a−2d) in d-Chloroform
a
b
1
−, not available. All of the isomers 2a−2d showed identical chemical shifts for H and 13C NMR.
Identification of Metabolites 3−7. The chemical
DISCUSSION
■
structures of 3−5 and 7 were confirmed by HPLC and 2D-
TLC cochromoatographies with the synthetic standards. For
the glucose conjugate of 6, the corresponding HPLC fraction
was isolated and then incubated with cellulase or β-glucosidase.
A new peak produced by the enzymatic hydrolysis was
identified as 6 by the HPLC and 2D-TLC cochromatographies
with the reference standard. Interestingly, the enzymatically
liberated aglycone 6 predominantly consisted from the single
isomer observed at the retention time of 20.9 min, the reference
standard of which has four isomers in total.
Significant loss of the radiocarbon, that is, maximum 25.6%AR
within 2 days, was observed after the foliar application of 1-
RTZ and 1-RTE onto the cabbage plant (Table 1), which
indicated their immediate volatilization from the leaf surface
due to the high vapor pressure. The volatilization of pesticides
from leaf surfaces can be conveniently evaluated with Henry’s
law constant. A similar trend in volatility as for 1 (2.1 × 10−4
m3 mol−1) was observed for other pyrethroids possessing a high
Henry’s law constant such as fenpropathrin (1.8 × 10−4 m3
mol−1), phenothrin (1.4 × 10−6 m3 mol−1), permethrin (1.9 ×
2612
dx.doi.org/10.1021/jf203903r | J. Agric. Food Chem. 2012, 60, 2607−2616

Journal of Agricultural and Food Chemistry
Article
1
Figure 3. H−1H COSY spectra of 2a and 1-RTZ in d-chloroform (273 K): (A) spectrum of 1-RTZ; (B) spectrum of metabolite 2a.
Figure 4. 1D-NOE spectra of 2 isomers in d-chloroform (273 K). (A, B) 1D-NOE spectra (5000 scans, delay = 5.0 s, τm = 0.375 s, LB = 0.5 Hz) of
2a or 2b (left) and mixture of 2c and 2d (right) after selective irradiation of proton at the ozonide ring. The irradiation targets are indicated by a tilde
(∼). The characteristic NOE response between the protons at the ozonide ring was observed only for the mixture of 2c and 2d by simultaneous
irradiation of the proton signals at the ozonide ring (signals marked by an asterisk). (C) Conventional 1H spectrum (16 scans) of 2a or 2b (left) and
mixture of 2c and 2d (right).
10−6 m3 mol−1), and cypermethrin (4.2 × 10−7 m3 mol−1),
on the leaf surface for a relatively longer period, such as
whereas those with a low hHenry's law constant tend to remain
tralomethrin (3.9 × 10−15 m3 mol−1).8−13
2613
dx.doi.org/10.1021/jf203903r | J. Agric. Food Chem. 2012, 60, 2607−2616

Journal of Agricultural and Food Chemistry
Article
Figure 5. Scheme of diastereomeric secondary ozonide production via the Criegee intermediate.
Both E/Z geometrical isomers reacted with atmospheric O3
on the leaf surface to form diastereomeric 2 as the major initial
degradates. There are many examples indicating the possible
formation of ozonide formation as a degradation intermediate
of pyrethroids, but the detection of ozonide is limited due to
their instability. Ruzo et al. studied the ozonolysis of synthetic
pyrethroids, phenothrin and halothrin, in several organic
solvents or on thin films and detected their ozonide derivatives,
which were tentatively identified by chemical ionization−mass
bulky substituents, is that trans alkenes are more likely to be
transformed to trans secondary ozonide via trans primary
ozonide and syn carbonyl oxide, and cis alkenes, vice
versa.15,17−19 The production of four diastereomers in the
ozonolysis of olefins can be explained from a generalized
observation on the “Criegee intermediate”,17−22 as summarized
in Figure 5, which can be briefly expressed by three steps: (1)
1,3-dipolar cycloaddition of O3 to olefin to produce a highly
unstable primary ozonide, (2) dissociation of the primary
ozonide to give the corresponding zwitterionic carbonyl oxide
and carbonyl compounds, that is, the Criegee intermediate, and
(3) recombination of the Criegee intermediate via another 1,3-
dipolar cycloaddition to produce more stable secondary
ozonide. Step 2 is considered to be a very important reaction
to produce four diastereomers on the cabbage leaf surface, in
which the four patterns of nonhomologous random recombi-
nation of carbonyl oxide ions and carbonyl compounds
produced from the primary ozonide may be deeply involved.
Incidentally, the secondary ozonides are considered to be less
toxic to mammals because they are highly likely to be unstable
in mammalian tissue due to their rapid decomposition in
aqueous solutions.23−25 Furthermore, an ozonide derivative of
unsaturated lipid, methyl oleate, was orally administered to rats,
which resulted in low toxicity, probably due to its fast
degradation by enzymes and biological antioxidants such as
glutathione transferase and vitamins, respectively.25
In the leaf tissue, the sugar conjugate of 6 was produced as a
major metabolite only in leaves treated with 1-RTZ. This may
be explained by the selective epoxidation of 1-RTZ to produce
intermediate 3 under the function of enzymes probably by
cytochrome P450 (CYP), because CYP-catalyzed stereo-
selective epoxidation of olefins is a common reaction in living
organisms such as lipid/hormone biosynthesis and xenobiotic
detoxification.26−29 Although 3 was detected from neither 1-
RTZ nor 1-RTE in cabbage plant, the epoxide derivative of 1 is
reported to be produced in rat and via photodegradation on
soil.3,30 Vaz et al. clarified that the epoxidation of olefins with
the CYPs from several origins proceeded more favorably with
(Z)-2-butene than its (E)-isomer.28 Sauveplane et al. reported
the CYP derived from Arabidopsis (Arabidopsis thaliana), which
was heterologously expressed in yeast (Saccharomyces cerevi-
siae), showed highly selective conversion of (Z)-configured
unsaturated fatty acids to mono- or di-cis-epoxide.29 Inciden-
tally, Ando et al. extensively studied the configuration of the
epoxy chrysanthemate derivatives of (S)-bioallethrin and
phenothrin produced by mouse microsomal CYP.31−33 They
1
spectrometry (CI-MS) and 19F, 13C, and H NMR spectros-
copy.14 Nambu et al. studied the metabolism of phenothrin on
bean foliage and identified its ozonide form by measuring its
molecular weight with electron ionization−mass spectrometry
(EI-MS) and detecting its typical degradates produced, such as
the carbonaldehyde and carboxylic acid derivative during the
1D-TLC development using an acidic solvent.8 Incidentally, the
halogen-substituted pyrethroids, for example, deltamethrin,
form lesser amounts of ozonide products14 and corresponding
degradates in the plant metabolism.9−11 Substituent groups at
olefins are considered to be an important factor to determine
the reactivity with ozone because an electron-withdrawing
substituent reduces the electron density of the olefin, which
suppresses the 1,3-dipolar cycloaddition of ozone, whereas an
electron-donating group enhances it.15 Because 2 has two
asymmetric carbons at the 1,2,4-trioxolane ring, there are four
diastereomers 2a−2d that could be separated by using HPLC
method 2. The synthetic reference 2 was confirmed to be
identical with those generated from 1-RTZ and 1-RTE on the
leaf surface of cabbage plant, and the isomeric ratio was also
similar between them. From the detailed analysis of the
exclusive data obtained from the 1D-NOE measurement
together with the spectrum pattern and chemical shifts/
coupling constants of various NMR analyses, the diastereomeric
configurations of 2 isomers were determined as trans (threo) for
2a and 2b and cis (erythro) for 2c and 2d. In the 13C NMR
spectra, the chemical shifts of C1 and C6 attached to the
ozonide ring shifted to lower field and C11 shifted to higher
field for 2a and 2b compared to those of 2c and 2d. This
difference in the chemical shifts was in good agreement with
the results reported by Griesbaum et al.,16 who conducted
extensive analyses to determine the configuration of the several
ozonide derivatives by 13C NMR analysis. Furthermore, 2a and
2b were more likely to be produced from 1-RTE, whereas 2c
and 2d were formed from 1-RTZ by ozonolysis. This result
supports our diastereomeric assignment; the empirical trend in
selectivity, which has been theoretically examined, especially for
2614
dx.doi.org/10.1021/jf203903r | J. Agric. Food Chem. 2012, 60, 2607−2616

Journal of Agricultural and Food Chemistry
Article
(8) Nambu, K.; Ohkawa, H.; Miyamoto, J. Metabolic fate of
phenothrin in plants and soils. J. Pestic. Sci. 1980, 5, 177−197.
(9) Mikami, N.; Baba, Y.; Katagi, T.; Miyamoto, J. Metabolism of the
synthetic pyrethroid fenpropathrin in plants. J. Agric. Food Chem. 1985,
33, 980−987.
(10) Cole, L. M.; Casida, J. E.; Ruzo, L. O. Comparative degradation
of the pyrethroids toralomethrin, tralocythrin, deltamethrin, and
cypermethrin on cotton and bean foliage. J. Agric. Food Chem. 1982,
30, 916−920.
(11) Gaughan, L. C.; Casida, J. E. Degradation of trans- and cis-
permethrin on cotton and bean plants. J. Agric. Food Chem. 1978, 26,
525−528.
(12) Agency for Toxic Substances and Disease Registry. Toxico-
logical profile for pyrethrin and pyrethroids, U.S. Department of
Health and Human Services, September 2003; available at http://
www.atsdr.cdc.gov/ToxProfiles/tp155.pdf.
have clarified that the oxygen atom was inserted into the
propenyl double bond from the si face to produce the 7,8-epoxy
derivative and its configuration at C7 carbon, the one adjacent
to the cyclopropane ring, was R and S from selective oxidation
of trans- and cis-chrysanthemates, respectively. Additionally,
epoxy hydrolases, which transform epoxy derivatives into diols,
are also reported to be stereoselective. Soybean-derived epoxy
hydrolase converts cis-epoxy fatty acids into threo diols by
catalyzing trans addition of water into oxirane carbon, which
has the S-chirality.34 Taking all above into consideration, a
single isomer of aglycone 6 detected in the cabbage plant is
most likely to be the threo isomer in R−R configuration, which
was produced via the intermediate cis isomer of 3 in R−S
configuration, as its estimated pathway is proposed in Figure 6.
In summary, the proposed metabolic pathway of 1 in the
cabbage plant is described in Figure 1. 1-RTZ and 1-RTE
(13) Laskowski, D. A. Physical and chemical properties of
pyrethroids. Rev. Envion. Contam. Toxicol. 2002, 174, 49−170.
(14) Ruzo, L. O.; Kimmel, E. C.; Casida, J. E. Ozonides and epoxides
from ozonization of pyrethroids. J. Agric. Food Chem. 1986, 34, 937−
940.
(15) Pryor, A. W.; Giamalva, D.; Church, D. F. Kinetics of ozonation.
3. Substituent effects on the rates of reaction of alkenes. J. Am. Chem.
Soc. 1985, 107, 2793−2797.
(16) Griesbaum, K.; Quinkert, R. O.; McCullough, K. J. C−C bond
formation at ozonide rings by substitution of chlorinated ozonides.
Eur. J. Org. Chem. 2004, 17, 3657−3662.
(17) Bauld, N. L.; Thompson, J. A.; Hudson, C. E.; Bailey, P. S.
Stereospecificity in ozonide and cross-ozonide formation. J. Am. Chem.
Soc. 1968, 90, 1822−1830.
Figure 6. Proposed enzymatic dihydroxylation pathway of 1-RTZ.
(18) Ponec, R.; Yuzhakov, G.; Hass, Y.; Samuni, U. Theoretical
analysis of the stereoselectivity in the ozonolysis of olefins. Evidence
for a modified criegee mechanism. J. Org. Chem. 1997, 62, 2757−2762.
(19) Bailey, P. S.; Ferrell, T. M. Mechanism of ozonolysis. A more
flexible stereochemical concept. J. Am. Chem. Soc. 1978, 100, 899−905.
(20) Murray, R. W.; Youssefyeh, R. D.; Story, P. R. Ozonolysis. Steric
and stereochemical effects in the olefin. J. Am. Chem. Soc. 1967, 89,
2429−2434.
reacted with atomospheric O3 on the leaf surface to form
ozonide as the initial product and successively decomposed to
carbonaldehyde and carboxylic acid derivatives via opening of
the trioxolane ring. In the leaf cell, they were further converted
to the polar metabolites via cleavage of the ester linkage and
dihydroxylation at the olefinic carbon followed by conjugation
with sugar. The isomerization on/in the cabbage leaf was a
minor reaction.
(21) Criegee, R. Mechanism of ozonolysis. Angew. Chem. Int. Ed.
1975, 14, 745−752.
(22) Geletneky, C.; Berger, S. The mechanism of ozonolysis revised
by 17O-NMR spectroscopy. Eur. J. Org. Chem. 1998, 8, 1625−1627.
(23) Zaikov, G.; Rakovsky, S. Ozonation of Organic and Polymer
Compounds; Smithers Rapra: Shropshire, U.K., 2009; Chapter 3, pp
179−218.
AUTHOR INFORMATION
■
Corresponding Author
*E-mail: andod@sc.sumitomo-chem.co.jp.
(24) Perry, S. C.; Charman, A. S.; Prankerd, J. R.; Chiu, K. F.
Chemical kinetics and aqueous degradation pathways of a new class of
ozonide antimalarials. J. Pharm. Sci. 2006, 95, 737−747.
(25) Pryor, A. W. Can vitamin E protect humans against the
pathological effects of ozone in smog? Am. J. Clin. Nutr. 1991, 53,
702−722.
(26) Guengerich, F. P. Cytochrome p450 oxidations in the
generation of reactive electrophiles: epoxidation and related reactions.
Arch. Biophys. Biochem. 2003, 409, 59−71.
(27) Hayashi, O.; Kameshiro, M.; Satoh, K. Intrinsic bioavailability of
14C-heptachlor to several plant species. J. Pestic. Sci. 2010, 35, 107−
113.
(28) Vaz, A. D. N.; McGinnity, D. F.; Coon, M. J. Epoxidation of
olefins by cytochrome P450: evidence from site-specific mutagenesis
for hydroperoxo-iron as an electrophilic oxidant. Proc. Natl. Acad. Sci.
U.S.A. 1998, 95, 3555−3560.
(29) Sauveplane, V.; Kandel, S.; Kastner, P.; Ehlting, J.; Compagnon,
V.; Wrek, R. D.; Pinot, F. Arabidopsis thaliana CYP77A4 is the first
cytochrome P450 able to catalyze the epoxidation of free fatty acids in
plants. FEBS J. 2009, 276, 719−735.
REFERENCES
■
(1) Matsuo, N.; Ujihara, K.; Shono, Y.; Iwasaki, T.; Sugano, M.;
Yoshiyama, T.; Uwagawa, S. Discovery and development of a novel
pyrethroid insecticide ‘Metofluthrin (SumiOne®, Eminence®)’.
Sumitomo Kagaku II 2005, 1−15.
(2) Kodaka, R.; Suzuki, Y.; Sugano, T.; Katagi, T. Aerobic
metabolism and adsorption of pyrethroid insecticide metofluthrin in
soil. J. Pestic. Sci. 2007, 32, 393−401.
(3) Nishimura, H.; Suzuki, Y.; Nishiyama, M.; Fujisawa, T.; Katagi, T.
Photodegradation of insecticide metofluthrin on soil, clay minerals and
glass surface. J. Pestic. Sci. 2011, 36, 376−38.
(4) Nishiyama, M.; Suzuki, Y.; Katagi, T. Hydrolysis and photolysis of
insecticide metofluthrin in water. J. Pestic. Sci. 2010, 35, 447−455.
(5) Baker, E. A. Chemistry and morphology of plant epicuticular
waxes. In The Plant Cuticle; Cutler, D. F., Alvin, K. L., Price, C. E.,
Eds.; Academic Press: London, U.K., 1982; pp 139−166.
(6) Katagi, T. Photodegradation of pesticides on plant and soil
surfaces. Rev. Environ. Contam.Toxicol. 2004, 182, 1−195.
(7) Meier, U. Leaf vegetable (forming heads). In Growth Stages of
Mono- And Dicotyledonous Plants; BBCH Monograph; Federal
Biological Research Centre for Agriculture and Forestry: Berlin,
Germany, 2001; pp 120−123.
(30) Kaneko, H. Pyrethroid chemistry and metabolism. In Hayes’
Handbook of Pesticide Toxicology, 3rd ed.; Krieger, R., Ed.; Elsevier:
Amsterdam, The Netherlands, 2010; Chapter 76, pp 1635−1663.
2615
dx.doi.org/10.1021/jf203903r | J. Agric. Food Chem. 2012, 60, 2607−2616

Journal of Agricultural and Food Chemistry
Article
(31) Ando, T.; Toia, R. F.; Casida, J. E. Epoxy and hydroxy
derivatives of (S)-bioallethrin and pyrethrins I and II: synthesis and
metabolism. J. Agric. Food Chem. 1991, 39, 606−611.
(32) Ando, T.; Jacobsen, N. E.; Toia, R. F.; Casida., J. E.
Epoxychrysanthemates: two-dimensional NMR analyses and stereo-
chemical assignments. J. Agric. Food Chem. 1991, 39, 600−605.
(33) Class, T. J.; Ando, T.; Casida, J. E. Pyrethroid metabolism:
microsomal oxidase metabolism of (S)-bioallethrin and the six natural
pyrethrins. J. Agric. Food Chem. 1990, 38, 529−537.
(34) Blee, E.; Schuber, F. Regio- and enantioselectivity of soybean
fatty acid epoxide hydrolsase. J. Biol. Chem. 1992, 286, 5197−5203.
2616
dx.doi.org/10.1021/jf203903r | J. Agric. Food Chem. 2012, 60, 2607−2616